The present invention provides a high throughput screening assay useful for detecting the presence of an exogenous DNA sequence in a sample. The method of the present invention further includes a high throughput DNA extraction method useful for extracting DNA from avian blood for subsequent use in a screening assay as, for example, an assay to detect the insertion of foreign DNA in the genome of a recipient.
The publications cited herein to clarify the background of the invention and in particular, materials cited to provide additional details regarding the practice of the invention, are incorporated herein by reference, and for convenience are cited in the following text.
Transgenesis is the ability to introduce foreign or exogenous DNA into the genome of a recipient, as for example, into a sheep, a cow or even a chicken. The ability to alter the genome of an animal immediately suggests a number of commercial applications, including the production of an animal able to express an exogenous protein in a form that is harvested easily.
The main obstacle to avian transgenesis is the low efficiency of introduction of foreign DNA into the chicken genome. The insertion of foreign DNA into the chicken genome using procedures that have worked for other animals is a difficult task and attempts at such have been mostly unsuccessful, partly due to the unique physiology of the chicken (Love et al., Transgenic birds by DNA microinjection, Biotechnology 12: 60–63, 1994; Naito et al., Introduction of exogenous DNA into somatic and germ cells of chickens by microinjection into the germinal disc of fertilized ova, Mol Reprod Dev 37: 167–171, 1994).
Through the use of retroviruses, a number of research groups have successfully introduced foreign DNA into the chicken genome at acceptable but low efficiencies (Bosselman et al., Germline transmission of exogenous genes in the chicken, Science 243: 533–5, 1989; Petropoulos, et al., Appropriate in vivo expression of a muscle-specific promoter by using avian retroviral vectors for gene transfer [corrected] [published erratum appears in J. Virol 66: 5175, 1992] J. Virol 66: 3391–7, 1992; Thoraval, et al., Germline transmission of exogenous genes in chickens using helper-free ecotropic avian leukosis virus-based vectors, Transgence Res 4: 369–377, 1995). The retroviral vectors used have been engineered such that they will not result in the replication and spread of any new retroviruses. This allows production of transgenic chickens that are free of any retrovirus. However, because the retroviral vectors cannot propagate in the chicken, the transgene is not transmitted from cell to cell. Retroviral vectors are typically injected into the embryo of a freshly laid egg through a small window in the egg shell. Approximately 1% of the embryonic cells are transduced, such that one copy of the transgene is inserted into the cell's DNA. After sexual maturity and meiosis, 0.5% of sperm or oocytes carry the transgene. In order to obtain one transgenic bird, at least 200 chicks have to be screened. It is often desirable to obtain several transgenic chicks because different chromosomal insertions can lead to different levels of transgene expression. Thus, it is necessary to breed and screen hundreds to thousands of chicks, necessitating a method for high throughput genetic screening for detecting the desired genetic sequence.
Random chromosomal insertion of transgenes via non-retroviral methods has become the mainstay of transgenics in some domesticated animals including pigs, sheep, goats and cows. The primary method to introduce the transgene is the injection of linearized DNA containing the desired transgene into the pronucleus of a zygote. Up to 20% of Go offspring contain the transgene. The relative high efficiency of transgenesis offsets the high technical costs incurred during the procedure. Transgenes have been inserted into goats, for instance, that direct the expression of pharmaceuticals in mammary glands for subsequent secretion into milk (Ebert, et al., Transgenic production of a variant of human tissue-type plasminogen activator in goat milk: generation of transgenic goats and analysis of expression, Biotechnology 9: 835–8, 1991).
In chickens, injection of the zygote germinal disk has been accomplished but with limited success, in part due to additional complications associated with unique aspects of chicken physiology and embryogenesis (Love et al., 1994; Naito et al., 1994). One lab has successfully produced several transgenic chickens, which have incorporated the injected DNA into their chromosomes and passed the transgene on to offspring. Another lab attempted to reproduce the technique but failed. Zygote injections in chickens are difficult because the nucleus is very small and is about 50 microns below the yolk membrane. Thus, the DNA must be injected into the cytoplasm. As in mice, cytoplasmic injection of DNA results in inefficient incorporation of the transgene into the chromosomes. Chickens must be sacrificed in order to remove the zygote and each chicken yields only one zygote.
An important technical breakthrough was pioneered by Gibbins, Etches, and their colleagues at the University of Guelph by using blastodermal cells (BDCs) collected from embryonic stage X embryos at oviposition, e.g., the time when the egg is laid (Brazolot et al., Efficient transfection of chicken cells by lipofection, and introduction of transfected blastodermal cells into the embryo, Mol Reprod Dev 30: 304–12, 1991; Fraser, et al., Efficient incorporation of transfected blastodermal cells into chimeric chicken embryos, Int J Dev Biol 37: 381–5, 1993). Coupled with recent progress in the culturing of BDCs, which can still reconstitute the germline, the method theoretically enables random transgene addition via nonhomologous recombination as well as targeted gene engineering via homologous recombination.
At stage X, the embryonic blastoderm consists of 40,000 to 60,000 cells organized as a sheet (area pellucida) surrounded by the area opaca; it harbors presumptive primordial germ cells (PGCs) that have not yet differentiated into migrating PGCs. Dispersed BDCs can be transfected with an appropriate transgene and introduced into the subgerminal cavity of y-irradiated, recipient stage X embryos. Irradiation may selectively destroy presumptive PGCs and retard recipient embryo growth allowing injected cells additional time to populate the recipient blastoderm. Using genetic markers for feather color (black for Barred Rock and white for White Leghorn), Etches, Gibbins and their colleagues were able to show that, of injected embryos surviving to hatch, 50% or greater of these were somatic chimeras of which nearly half were also germline mosaics (Petitte, et al., Production of somatic and germline chimeras in the chicken by transfer of early blastodermal cells, Development 108: 185–9, 1990).
Gibbins and her colleagues have determined that random gene addition occurs in in vitro cultured BDCs in 1 out of every 300 transfected cells (Gibbins and Leu, personal communication). They did not determine whether BDCs with random gene additions can be re-introduced into stage X embryos to produce germline Go chimeras. Therefore, the actual efficiency of transgenesis has not yet been determined.
Gene targeting, the ability to specifically modify a specific gene, is a much sought-after technology in a variety of species, including chickens, because such modifications will result in very predictable transgene expression and function. Gene targeting has been successfully accomplished in mice because mouse embryonic stem (ES) cells can be cultured in vitro for long periods of time and still contribute to the germline (Mountford et al., Dicistronic targeting constructs: reporters and modifiers of mammalian gene expression, Proc Natl Acad Sci U.S.A. 91: 4303–7, 1994). The long-term culture of mouse ES cells allows the researcher to select for and expand colonies of cells transfected with the targeting vector that have the transgene inserted into the proper site. Similar to the use of the feather color alleles in chimeric birds, coat color of different breeds of mice are used to track the donor cells in offspring. The difficulty in applying the mouse ES cell technology to other species is that it has been impossible to isolate ES cells of other species. While cells resembling ES cells have been isolated from goats and pigs and cultured in vitro, these cells are not able to contribute to recipient embryos after long-term culture. Nuclear transfer technology offers an alternative to the use of ES cells and it is probable that gene targeting in animals will, in the future, be implemented via nuclear transfer. Presently, however, nuclear transfer is very inefficient and expensive, making its implementation a slow process.
Recent advances in the in vitro short-term culture of chicken blastodermal cells, combined with the unique physiology of avian reproduction, indicate that gene targeting is possible in chickens. The division rate of stage X BDCs can be maintained in vitro at one division every 8–10 hours for 4–8 days using culture conditions developed by the Ivarie laboratory (University of Georgia, Athens, Ga.) and AviGenics, Inc. (Athens, Ga.) (Speksnijder and Baugh, unpublished data). The ability to propogate BDCs in vitro at this rate, while maintaining totipotency, will allow for the rapid expansion of cell colonies containing the desired genetic modification. This, combined with the fact that large numbers of BDCs (40,000 to 60,000 cells/egg) can easily be isolated from freshly laid chicken eggs, makes it feasible to screen large numbers of transfected BDC colonies for those having a desired gene of interest.
Currently, BDCs can only be cultured for 4 to 8 days before they lose the ability to contribute to germ tissues in the recipient embryo (Speksnijder and Baugh, unpublished data). Therefore, it is likely that BDCs carrying the desired genetic modification can only be enriched to perhaps 0.1 to 10% of the total number of donor cells. While sufficient to enable gene targeting, the rate of transmission of the desired genetic modification from chimeric founder animals (those that were directly derived from injection of donor BDCs into recipient embryos) to their offspring will be low. Hundreds to thousands of offspring will have to be screened, again necessitating a method for high throughput genetic screening for detecting a desired sequence.
The enrichment of BDCs for desired genetic modifications can be applied to transgenesis projects involving random insertion of a gene into the avian genome, as well as modification of a specific gene. Therefore, a method for high throughput genetic screening will have broad applications in the fast-growing field of avian transgenesis.
To determine if an organism contains a novel or new gene, DNA is extracted from a tissue sample (blood, skin, sperm) and is subjected to an assay that will detect the gene. The method of choice was the Southern assay, which is extremely sensitive and reliable (Southern, E. M., Detection of specific sequences among DNA fragments separated by gel electrophoresis, J Mol Biol 98, 503–17, 1975). However, the Southern assay is very labor intensive and time consuming.
The Southern assay was replaced by the polymerase chain reaction (PCR) method (Mullis et al., Specific enzymatic amplification of DNA in vitro: the polymerase chain reaction. Cold Spring Harbor Symp Quant Biol 51 (Pt 1): 263–73, 1986), which is a more sensitive and rapid assay.
Recently developed techniques, such as the TAQMAN sequence detection system (Applied Biosystems, Foster City, Calif.) allow hundreds of samples to be analyzed in hours without requiring a time-consuming gel electrophoresis step (Heid et al., Real time quantitative PCR, Genome Res 6: 986–94, 1996). During a TAQMAN reaction run, which is setup like a PCR reaction, a fluorogenic probe consisting of an oligonucleotide with both a reporter and a quencher fluorescent dye attached, anneals specifically between the forward and reverse primers. The probe and primers are complementary to the sequence of the desired transgene. When the probe is cleaved by the 5′ nuclease activity of Taq DNA polymerase, the reporter dye is separated from the quencher dye and a sequence-specific signal is generated. With each cycle, additional reporter dye molecules are cleaved from their respective probes, and the fluorescence intensity is monitored during the PCR. Samples are analyzed in 96-well plates and, at the end of a run, it is obvious which samples contain the desired sequence.
While high throughput methods for sequence detection are available, no comparable methods exist for the extraction of DNA useful in a high throughput assay for sequence detection. Rather, existing DNA extraction methods are still labor intensive and time consuming. The majority of extraction methods require the DNA samples to be treated in individual tubes. Samples are subjected to a number of steps, including proteinase digestion, extraction with organic solvents, and precipitation. The extraction step is particularly problematic because of the awkwardness of manipulation of the solution phases. Salting out has been used as an alternative for extraction of unwanted proteins, but this method requires multiple centrifugations and tube transfers. Kits are available which avoid the extraction steps by using DNA binding resins and allow for the processing of 96 samples at a time. However, the resins are not reusable, and their use can result in poor yield and inconsistent DNA quality. In addition, these kits are not cost-effective, costing up to $3.00 per sample processed for extraction.
Existing methods for extracting DNA extraction from multiple samples of avian tissue are labor intensive and tedious. Avian blood, like all non-mammalian vertebrates, has a special quality in that the erythrocytes are nucleated (Rowley and Ratcliffe, Vertebrate blood cells, Cambridge University Press, Cambridge, N.Y., 1988). The presence of nucleated cells allows one to extract a large amount of DNA from a very small amount of blood. But existing DNA extraction techniques have not taken advantage of this aspect of avian blood. Grimberg, et al. developed a method in which the plasma membrane, but not the nuclear membrane, of red blood cells (RBCs) was lysed (Grimberg et al., A simple and efficient non-organic procedure for the isolation of genomic DNA from blood, Nucleic Acids Res 17: 8390, 1989). Subsequently, Petitte et al. augmented Grimberg's method by optimizing the initial lysis and spooling ethanol-precipitated DNA out on a glass rod, resulting in a more pure DNA preparation but requiring a more labor-intensive protocol (Petitte, et al., Isolation of genomic DNA from avian whole blood, Biotechniques 17: 664–6, 1994). Thoraval, et al. used a similar procedure, however, each sample was required to be treated individually (Thoraval et al., Germline transmission of exogenous genes in chickens using helper-free ecotropic avian leukosis virus-based vectors, Transgenic Res 4: 369–377, 1995).
All of the aforementioned procedures possess similar disadvantages in that the each sample must be treated individually and the DNA extracted must be transferred between multiple tubes. In addition to being labor-intensive, these DNA extraction procedures include an overnight incubation for lysis to occur.
In order to target genes in mice, hundreds of mouse embryonic stem (ES) cell colonies have to be individually analyzed for the presence of the desired genetic modification. In order to facilitate DNA extraction from a large number of colonies, Ramirez-Solis et al. devised an ingenious method in which ES cells are lysed in 96-well plates (Ramirez-Solis et al., Genomic DNA microextraction: a method to screen numerous samples. Anal Biochem 201: 331–5, 1992). Using the method of Ramirez-Solis, et al., DNA is precipitated such that it sticks to the bottom of the microtiter well without centrifugation. This is due in part to the affinity of DNA for polystyrene, the major component of 96-well tissue culture plates. While the DNA is stuck to the plates, all the unwanted protein and salts can be removed by washing the wells multiple times with 70% ethanol. In this way, 96 samples can be processed simultaneously. Because the DNA is not transferred among tubes, the possibility of both sample loss and contamination,is minimized.
Ramirez-Solis et al. attempted to isolate DNA from human blood samples using the above-described method, however the inefficiency of the procedure required processing a large volume of blood to obtain enough cells for efficient extraction. At least 0.3 ml, and most probably about 1.0 ml, of human blood is required per well to obtain enough DNA for efficient extraction, however the maximum capacity of each microtiter well is only about 0.25 ml. Thus, the method of Ramirez-Solis, et al. is not useful for the high throughput extraction of DNA from genomic blood.
Udy and Evans developed a 96-well plate method for DNA extraction from embryonic stem (ES) cells, similar to the method of Ramirez-Solis et al., but never applied their method to the extraction of DNA from blood (Udy and Evans, Microplate DNA preparation, PCR screening and cell freezing for gene targeting in embryonic stem cells, Biotechniques 17: 887–94, 1994).
In view of the aforementioned deficiencies of the prior art, there is a need for a method providing for the rapid and easy extraction of DNA from a large number of blood samples without necessitating large sample volumes, requiring the transfer of DNA between multiple tubes, or necessitating overnight incubation steps. Further, there is a need for a DNA extraction method that can be used in an high throughput assay to rapidly screen a large number of samples to detect a desired DNA sequence or transgene. Finally, there is a need for a high-throughput assay useful for detecting the presence of a desired genetic sequence in a large number of samples when the copy number is low, i.e., between about 5 to about 50 copies.